Decoding of genetic information into protein sequences is a multi-step process that requires specific charging of transfer RNA (tRNA) molecules with their cognate amino acids by their cognate aminoacyl-tRNA synthetases (aaRSs) to form aminoacyl-tRNA (aa-tRNA), specific binding of aa-tRNAs to the ribosomal decoding center programmed with their cognate triplet codons, and large scale movements of tRNAs within the ribosome as they shift progressively from the tRNA entry site (the A-site), to the peptidyl-tRNA site (the P-site), and finally to the tRNA exit site (the E-site), from which tRNA dissociation takes place.
While the elementary steps in elongation have been identified, additional studies will be required to achieve a full understanding of the molecular mechanisms of each step. Of particular interest is the role of tRNAs, which, rather than being passive and rigid substrates for the ribosome, have been more recently implicated as being “active” players in the decoding process (Westhof, 2006), interacting strongly with different sites on the ribosome and undergoing substantial conformational changes in migrating from the A- to P- to E-sites (Korostelev et al., 2006; Selmer et al., 2006). For example, a mutation in the D stem of tRNATrp (known as the Hirsh suppressor mutation (Hirsh, 1971) has been shown to promote miscoding at the anticodon. Recent kinetic studies show that this mutation specifically accelerates two forward steps on the ribosome: GTP hydrolysis for accommodation of aa-tRNA to the A site and peptide bond formation (Cochella & Green, 2005). Because this mutation is distal from the codon-anticodon interaction, its ability to promote tRNA accommodation and peptide bond synthesis suggests that the tRNA body is in direct communication with both the decoding center of the 30S subunit and the GTPase center of the 50S subunit. Another example is provided by the tertiary core ‘elbow’ region of tRNA, which is formed by extensive interactions between the D and T-loops. We have demonstrated, using single turnover rapid kinetics measurements (Pan et al., 2006; Pan et al., 2007), that mutations in the conserved G18:U55 base pair interfere with the ribosomal translocation step, particularly for tRNA moving from the P- to the E-site, consistent with X-ray crystallography results showing that position 55 is in direct contact with protein L1 in the E-site (Korostelev et al., 2006).
In existing studies, an important assay for the translocation rate is based on fluorescent changes of modified natural tRNAs, isolated from E. coli or yeast cells, whose D residues have been replaced with proflavin (Wintermeyer & Zachau, 1979; Savelsbergh et al., 2003). However, this approach, as so far applied, suffers from two significant limitations. First, it does not allow direct monitoring of the movements of tRNA mutants on the ribosome. This is because mutant tRNAs, which are prepared by run-off in vitro transcription with T7 RNA polymerase (Sampson et al., 1989), lack D residues. As a result fluorescent A-site tRNA has been used to monitor effects of mutation in P-site tRNA, and fluorescent P-site tRNA to measure effects of mutation in A-site tRNA. Second, proflavin is rapidly photobleached, rendering proflavin-labeled tRNA unsuitable for single molecule experiments in which fluorescent probes are subject to high light fluxes. Interest in overcoming this limitation is high, because recent work has clearly demonstrated the potential of the single-molecule approach to yield more detailed mechanistic information about protein synthesis than is available from ensemble single turnover experiments (Blanchard et al., 2004a; Blanchard et al., 2004b).
The labeling of tRNA has been performed with respect to each of four different components of such molecules: (1) Amino acids. The amino group of Lys-tRNAlys was labeled with BODIPY FL by displacing its succinimidyl group (Woodhead, 2004), and the amino group of Met-tRNAfMet was acylated and reacted with maleimide to produce fluorophore-Met-tRNAfMet, which, however, has reduced activity compared to the unmodified molecule (McIntosh 2000). (2) 4-thioU(8) group. This has been used for the studies of aminoacyl-tRNA binding to the ribosomal A site (Bieling et al, 2006; Blanchard, 2004a,b; Munro, 2007). (3) acp3U47 group. It has only been labeled by the Blanchard group for their FRET studies (Blanchard, 2004a,b; Munro, 2007). (4) Dihydrouridine group. This group has been used to study the kinetics of tRNA binding and movements on the ribosome, but the choice of dye is limited to only proflavin, which, as described above, is very sensitive to environment but is of little use for single molecule studies, which require brighter dyes, or for FRET studies, which require a good donor-acceptor pair.
Since labeled tRNAs are so important for the studies of dynamics of ribosome function it is important to find a universal method of labeling many tRNA species with many different dyes. All the above labeling methods have their limitations. The amine reactive labeling of the amino acid results in tRNAs that have low activities (McIntosh, 2000; Woodhead, 2004). The other three methods are dependent on the existence of the modified group, so they can not be used if a transcribed tRNA, e.g., for the in vitro study of tRNA mutants, is required. For acp3U47, only 5 of the 20 amino acids have tRNAs that have a acp3 modification further limiting its application on other tRNAs. 4-thioU and dihydroU are more prevalent modifications occurring in the 8th position, and D loop, respectively (in E. coli only tRNAGlu, tRNALys, and tRNAThr do not have a 4-thioU modification, and only tRNAGlu and tRNATyr do not have a D modification). In ribosome studies, 4-thio U has been only successfully used with initiator tRNA, and our attempts to label an elongator tRNA proceeded with only modest yields. Furthermore, labeled Tyr-tRNATyr showed very poor binding to ribosomes under conditions that are EF-Tu dependent.
Over the last decade, achievements have been realized through the application of new technologies to the life sciences, for example, whole genome sequencing, DNA microarrays, and proteomic high-throughput analysis. The data obtained with these technologies serve to underscore gaps remaining in the cellular information currently available. Two of these gaps are: 1) sensitive and efficient protein identification, and 2) the dynamics of protein expression. Development of methods to detect protein synthesis directly and in real time, identify the amino acid sequence of a protein, and localize such synthesis within a cell (live proteomics) will enable fundamental advances in understanding basic life processes and aid significantly the search for new sources of therapy. See, for example, PCT Published Apps. WO 2004/050825 and WO 2006/228708. Protein synthesis monitoring (PSM) is an analytical method to identify proteins being synthesized on single ribosomes, in live cells, and in real time. In PSM, the protein synthesis apparatus is marked with a unique fluorescent labeling scheme, producing sequence-specific signals that enable protein identification. See, for example, PCT Published App. WO 2005/116252.
The study of cellular dynamics can utilize mRNA profiling in tissues and cells as a primary tool in research and clinical diagnosis. However, mRNA levels are uncertain predictors of protein expression. Current proteomic analysis, based largely on 2D electrophoretic gels, mass spectrometry, and combinatorial arrays, is limited by destructive sample preparation, preventing both the real-time detection of proteins and the elucidation of the dynamics of cellular response to various modulators. A need exists in the art for reagents with increased sensitivity in a system for studying cellular dynamics, such as Protein Synthesis Monitoring (PSM), which can measure protein synthesis by following thousands of labeled ribosomes simultaneously and repeating the measurements at any specific location for hours or days. Fluorescently translation components with increased sensitivity in a protein synthesis monitoring system are needed to provide the ability to record the dynamic patterns of protein synthesis in live cells in vivo, or in vitro.